anaerobic bacterium Desulfococcus biacutus
Dissertation submitted for the degree of Doctor of Natural Sciences
Presented by
Olga Brígida Gutiérrez Acosta
at the
University of Konstanz
Faculty of Sciences Department of Biology
Date of the oral examination: 06.december.2013 First supervisor: Prof. Dr. Bernhard Schink Second supervisor: Prof. Dr. Jörg Hartig Third supervisor: Prof. Dr. Alasdair Cook
Konstanzer Online-Publikations-System (KOPS) URL: http://nbn-resolving.de/urn:nbn:de:bsz:352-267797
doctoral research work. Thanks for giving me the opportunity to develop my doctoral thesis in your working group, for guiding me, for sharing ideas, and for all discussions.
I thank the members of my thesis committee Prof. Alasdair Cook, Prof. Jörg Hartig and Prof. Bernhard Schink, for their participation in my evaluations as a member of the graduate school Konstanz Research School Chemical Biology. Thank you for the pleasant discussions and valuable comments.
I also thank Prof. Peter Kroneck for the valuable discussions about my work, and for the personal support. I am also thankful to Dr. David Schleheck, Dr. Felix Ten Brink and Dr. Diliana Simeonova for their help and discussion.
Thanks to all members and participants of the DFG priority program 1319, for recognizing the scientific advance made on this theme during my doctoral research. I especially thank Prof. Bernard T. Golding for the valuable discussions about my work.
I am grateful to all members of the AG Schink for the help and collaborations during my doctoral research and for the coffee and cake time.
My warm thanks to all my friends for the great moments during my stay in Konstanz, with special thanks to Natalia Charlina and to Jennifer Ignatious for the sincere and lovely friendship.
My most profound thank to my parents, brothers, and all the new members of the Gutierrez Acosta family for their support in this personal achievement. Immensely thanks to my husband Norman Hardt for his encouragement, professional support and companionship.
agradecimientos a mi supervisor de tesis, el Prof. Bernhard Schink. Gracias por darme la oportunidad de desarrollar mi tesis en su grupo de trabajo y por guiarme acertadamente durante la realización de la misma.
Agradezco a los miembros de mi comité de tesis Prof. Alasdair Cook, Prof. Jörg Hartig y Prof. Bernhard Schink, por haber participado como tales durante mis evaluaciones como miembro de la escuela de graduados Konstanz Research School Chemical Biology. Gracias por las gratas discusiones y sugerencias que me ayudaron a tener un mejor desempeño en mi trabajo.
Quiero agradecer también al Prof. Peter Kroneck por las discusiones sobre mi trabajo, y por el apoyo personal. Así mismo agradezco al Dr. David Schleheck, Dr.
Felix Ten Brink y Dr. Diliana Simeonova por sus sugerencias y apoyo en la realización de mi trabajo.
Gracias a los miembros y participantes del programa científico alemán de prioridad DFG 1319 por reconocer el avance científico en el tema, logrado durante mi tesis doctoral. Agradezco especialmente al Prof. Bernard T. Golding por sus apreciadas discusiones sobre mi trabajo.
A todos los miembros del grupo de trabajo del Prof. Schink, por su apoyo y colaboración durante mi estancia doctoral y por todos los momentos de café.
Gracias a todos mi amigos que me brindaron gratos momentos durante mi estancia en Konstanz. En especial muchas gracias a Natalia Charlina y Jennifer Ignatious por brindarme una amistad sincera.
Mis más profundos agradecimientos van hacia mis padres, mis hermanos y todos los nuevos integrantes de la familia Gutiérrez Acosta por apoyarme en este logro más en mi vida. Agradezco inmensamente a mi esposo Norman Hardt por el apoyo brindado tanto personal como profesional y por su compañía.
Table of Contents
CHAPTER 1 General Introduction………...…………...………1
1.1 Important aspects about acetone……….……1
1.2 Acetone degradation by aerobic bacteria……….1
1.3 Acetone degradation by nitrate-reducing bacteria………2
1.4 Acetone degradation by sulfate-reducing bacteria (SRB)………..3
1.5 Acetone degradation by fermenting bacteria………4
1.6 Energetic considerations of acetone degradation……….5
1.7 SRB and genome sequences………6
1.8 Hypothesis………..6
1.9 Aim of the thesis……….7
CHAPTER 2 Carbonylation as a key reaction in anaerobic acetone activation by Desulfococcus biacutus………...………...8
2.1 Abstract………...8
2.2 Introduction………9
2.3 Experimental procedures………...11
2.4 Results………...16
2.5 Discussion……….25
Acknowledgements………..28
CHAPTER 3 ATP and thiamine pyrophosphate dependence of acetone degradation by the sulfate-reducing bacterium Desulfococcus biacutus monitored by a fluorogenic ATP analogue………..29
3.1 Abstract………29
3.2 Introduction………..30
3.3 Materials and Methods………..………32
3.4 Results and Discussion………..34
3.5 Conclusions………39
Acknowledgements………40
CHAPTER 4 Acetone utilization under sulfate-reducing conditions: draft genome sequence of Desulfococcus biacutus and a proteomic
survey of acetone-inducible proteins………..…...…..…41
4.1 Abstract………41
4.2 Introduction………..41
4.3 Materials and Methods………..43
4.4 Results………...46
4.5 Discussion……….67
Acknowledgements………..……….69
CHAPTER 5 General Discussion………...………70
5.1 Activation of acetone by Desulfococcus biacutus……….70
5.2 Comparison with other hydrocarbon activation……….74
5.3 Future research………..76
SUMMARY………...77
ZUSAMMENFASSUNG……….78
RECORD OF ACHIEVEMENT………...79
ABGRENZUNG DER EIGENLEISTUNG……….80
REFERENCES……….………...81
SUPPLEMENTARY DATA………...91
SCIENTIFIC CONTRIBUTIONS LIST………..95
1
CHAPTER 1
General Introduction
1.1 Important aspects about acetone
Acetone production is an important process in synthetic chemistry. Due to its physicochemical properties, acetone represents one the most used solvents in industry.
Acetone is used mainly as an intermediate in the synthesis of methacrylate, bisphenol A, diacetone alcohol, and some other compounds (Sifniades et al., 2011). Acetone regulation was exempted by the EPA in 1995. It is neither considered as hazardous pollutant in the Clean Air Act, nor as a priority pollutant in the Clean Water Act (Sifniades et al., 2011).
However, exposure to high concentration of acetone vapors causes eye irritation and narcosis. In addition, acetone has been reported as a toxic compound (Singh et al., 1994).
Acetone gets into the environment also by bacterial fermentations, for example by several Clostridium species (Duerre et al., 1992; Han et al., 2011; Lépiz-Aguilar et al., 2013). It is also one of the three ketone bodies that are formed in the human body and are excreted by diabetic mammalians (Asagoe et al., 1968; Kalapos, 1997; Laffel, 1999). Therefore, the metabolic pathways by which acetone is degraded under different environments are of special interest.
1.2 Acetone degradation by aerobic bacteria
Acetone can be degraded by some aerobic bacteria and mammals (Bondoc et al., 1999;
Koop and Casazza, 1985; Park et al., 1995) via oxygen-dependent hydroxylation to acetol mediated by cytochrome P450, as it was shown with Mycobacterium smegmatis (Landau and Brunengraber, 1987; Taylor et al., 1980). However, an enzyme converting acetone to acetol has not been described yet. The first report on different ways of methyl ketones
2
degradation was made for hydrocarbon-utilizing bacteria (Lukins and Foster, 1963). The mechanism of acetone activation has been under discussion for several years. Evidence of a carboxylation reaction was shown with the photosynthetic bacterium Rhodopseudomonas gelatinosa (Siegel, 1950; Siegel, 1954). Requirement of CO2 as a co- substrate for acetone degradation was observed for first time with a methanogenic co- culture (Platen and Schink, 1987), with Thiosphaera pantotropha which was able to grow aerobically as well as anaerobically (Bonnet-Smits et al., 1988), and with Rhodobacter capsulatus and other phototrophs (Birks and Kelly, 1997). The need for CO2 suggested a carboxylation of the methyl group of acetone, forming acetoacetate. A CO2- and ATP- dependent activation of acetone was also observed with cell-free extracts of the aerobic bacterium Xanthobacter autotrophicus strain Py2 (Sluis and Ensign, 1997). The carboxylation of acetone was also investigated with Rhodococcus rhodochrous, but in this case the activity was not stimulated with ATP, but it depended on the presence of other nucleotides like GTP, ITP, CTP and UTP (Clark and Ensign, 1999).
Acetone carboxylase was first purified from Xanthobacter autotrophicus strain Py2 (Sluis and Ensign, 1997). Its further characterization and comparison with the carboxylase of the phototrophic bacterium Rhodobacter capsulatus showed that they are identical in subunit composition (α2β2γ2 multimers of 85-, 78-, and 20-kDa subunits) and in kinetic properties (Sluis et al., 2002). Electron paramagnetic resonance (EPR) spectra of cell extracts of acetone-grown cells of R. capsulatus showed that in this bacterium the acetone carboxylase was manganese-dependent (Boyd et al., 2004).
1.3 Acetone degradation by nitrate-reducing bacteria
Nitrate-dependent acetone oxidation was shown with Thiosphaera pantotropha (Paracoccus pantotrophus) as mentioned above. The degradation of acetone was also studied with the denitrifying bacterium strain BunN, which essentially required CO2 for growth with acetone (Platen and Schink, 1989). Therefore, it was concluded that the degradation of acetone by strain BunN occurs similar to T. pantotropha via carboxylation reaction. This initial reaction was proven with cell-free extracts of strain BunN, in which a decarboxylation of acetoacetate occurred in the presence of ADP and MgCl2, and was specific for acetone-grown cells (Platen and Schink, 1990). Since a carboxylation reaction
3
could not be proven, a different approach to support the carboxylation reaction was done with labeling experiments. Cell-free extracts of strain BunN catalyzed the exchange of
14CO2 into acetoacetate in an ADP-dependent reaction (Janssen and Schink, 1995a).
Acetoacetate was then converted to a labeled acetoacetyl-CoA by succinyl-CoA:
acetoacetate CoA transferase. Those experiments led to the conclusion that the activating reaction occurred in an ATP-dependent carboxylation of acetone. The carboxylating enzymes were analyzed with the nitrate reducers Alicycliphilus denitrificans K601, Paracoccus denitrificans, and Paracoccus pantotrophus (Dullius et al., 2011). The activity of carboxylase from the enriched enzymes was dependent on ATP, and could be detected also with butanone. However, the carboxylation reaction was also detected with UTP, ITP, and GTP. The enriched fractions were analyzed by SDS-PAGE where it was observed molecular weights similar to the aerobic acetone carboxylases. Sequence analysis of the enriched fractions resulted in similar subunit composition (85.3-, 78.3-, and 19.6-KDa) as it was found for the acetone carboxylases of X. autotrophicus and R. capsulatus. Recently, an ATP-dependent carboxylation of acetone was again shown with the nitrate reducer Aromatoleum aromaticum (Schühle and Heider, 2012) which was reported before to be able to carboxylate acetophenone to benzoylacetate during anaerobic ethylbenzene degradation (Jobst et al., 2010). In both carboxylation reactions the presence of ATP was needed. At the moment it is well established that aerobic and nitrate-reducing bacteria employ similar carboxylating mechanisms to activate acetone, and that the carboxylating enzymes are molecularly highly similar.
1.4 Acetone degradation by sulfate-reducing bacteria (SRB)
Under sulfate reducing conditions the degradation of acetone has been studied with the Gram-negative bacterium Desulfococcus biacutus (Janssen and Schnik, 1995; Platen et al., 1990), and with Desulfobacterium cetonicum (Janssen and Schink, 1995b). The degradation of acetone by SRB appeared to depend on the presence of CO2 as well. However, neither activity of acetone carboxylase was detected in cell-free extracts, nor acetoacetate- decarboxylating activity could be found. These results indicated that acetoacetate is not an intermediate in the activation of acetone by SRB. Formation of a free intermediate was excluded, based on the absence of CoA transferase or CoA ligase activity in cell-free
4
extracts of acetone-grown cells. The presence of acetoacetyl-CoA thiolase suggested that acetone may be activated in a reaction that probably leads directly to acetoacetyl-CoA.
The possibility of a carbonylation reaction leading to a 3-hydroxybutyryl-CoA derivative was studied with D. biacutus, but without any success (Dullius, 2011). Since D. biacutus oxidizes acetyl residues through the Wood-Ljungdahl pathway, it possesses CO dehydrogenase activity. Therefore, this bacterium is able to convert CO2 to CO and employ this as a co-substrate in acetone activation. Studies in cell-free extracts of acetone-grown cells did not show activity of 3-hydroxybutyryl-CoA dehydrogenase.
Moreover, 3-hydroxybutyrate was never found as the product of acetone activation.
These results indicated that 3-hydroxybutyrate is not an intermediate in the metabolism of acetone. From all these studies it was concluded that the activation of acetone in SRB, or at least in D. biacutus may occur through a mechanism different from the carboxylation reaction described for aerobic and nitrate-reducing bacteria.
1.5 Acetone degradation by fermenting bacteria
Acetone can also be degraded in the absence of electron acceptors (Platen et al., 1994;
Platen and Schink, 1987; Symons and Buswell, 1933; Wikén, 1940). An enrichment culture (WoAct) that was obtained from anoxic sediment was able to degrade acetone anaerobically (Platen and Schink, 1987). Microscopy studies showed that one of the cooperation partners was a filamentous bacterium highly similar to Methanothrix sp (Methanosaeta sp). Inhibition experiments with streptomycin to avoid acetone degradation, and with acetylene and bromoethanesulfonate to inhibit acetate degradation, confirmed that acetone was converted to acetate which was further degraded to methane and CO2. Further studies with the enrichment culture WoAct indicated that acetone degradation by the fermenting bacterium does not necessarily depend on acetate removal by the methanogenic partner (Platen et al., 1994). However, acetone degradation was impeded when the concentration of acetate reached 10 mM. Up to now it is clear that acetone can be degraded in syntrophy, but the mechanism of acetone activation in this process has not been studied in detail yet.
5
1.6 Energetic considerations of acetone degradation
Acetone is activated by aerobic and nitrate-reducing bacteria by carboxylation in a CO2- and ATP-dependent reaction to form acetoacetate. According to equation 1 (Eq. 1), activation of acetone via carboxylation is an endergonic reaction that requires less than one ATP.
(Eq. 1) CH3COCH3+CO2 →CH3COCH2COO−+H+ (∆G0´ = +17.1 kJ mol-1)
In the proposed mechanism, the γ and β phosphodiester bonds of ATP need to be hydrolyzed during the reaction; one is invested for the enolization of acetone (to the form of phosphoenolacetone), and the other one is used to carboxylate the enolized acetone to acetoacetate. After carboxylation, acetoacetate is activated to acetoacetyl-CoA with the investment of one further ATP. In total, two ATP equivalents are invested for acetone activation and one more for further acetoacetate degradation. This energy expenditure can be afforded by aerobic and nitrate-reducing bacteria because the subsequent oxidation of the acetyl moieties releases sufficient energy. Therefore, growth with acetone by those bacteria is possible not affected despite such energy expensive activation reaction.
Acetone degradation by SRB is energetically more difficult. According to equation 2, the degradation of acetone coupled to the reduction of sulfate yields less than two ATP.
(Eq. 2) CH COCH SO H CO2(aq) H2S(aq) H2O 2
4 3
3 +2 −+4 + →3 +2 +3 (∆G0´ = - 115.3 kJ mol-1)
Oxidation of the acetyl residue of acetyl-CoA through the CO dehydrogenase (“Wood- Ljungdahl”) pathway can form only about one ATP equivalent per acetyl residue. Thus, acetone degradation through the carboxylation reaction described above (eq. 1) could not be supported through the subsequent oxidation of the acetyl residues. Therefore, a different mechanism for CO2-dependent acetone activation has to be postulated for these bacteria.
6 1.7 SRB and genome sequences
SRB are prokaryotes that are able to perform dissimilatory sulfate reduction. Those prokaryotic microorganisms including bacteria and archaea are organized in five different phylogenetic lineages (Thauer et al., 2007): the mesophilic deltaproteobacteria, the Gram- positive bacteria, the thermophilic Gram-negative bacteria, the Euryarchaeota, and theThermodesulfobiaceae. SRB are ubiquitous in nature, therefore, they can be found in various anoxic environments where sulfate is present, like soil, sediments, marine and freshwater, for example. Depending on their ability to oxidize the C2 carbon unit of acetyl-CoA to CO2, they can be grouped as complete oxidizers or incomplete oxidizers.
There are two known ways for the oxidization of the acetyl residues; the first one is through the tricarboxylic acid cycle, and the second one is through the carbon monoxide dehydrogenase (CODH) pathway.
The completed genomes of several SBR are already available. Among them are found:
Archaeoglobus sulfaticallidus strain PM70-1 (Stokke et al., 2013), the marine deltaproteobacterium Desulfobacula toluolica Tol2 which is able to degrade aromatic compounds (Wohlbrand et al., 2013), Desulfobacca acetoxidans strain ASRB2 (Goker et al., 2011), Strain NaphS2 which can grow anaerobically on naphthalene (DiDonato et al., 2010), Desulfobacterium autotrophicum strain HRM2 able to grow on fatty acids (Strittmatter et al., 2009), for example. Despite the sequenced genomes of SRB, the genome of an acetone-degrader sulfate reducing microorganism has not been reported until now.
1.8 Hypothesis
Since the degradation of acetone by D. biacutus theoretically cannot be driven by the same carboxylation mechanism that is employed by aerobic and nitrate reducers, a novel mechanism must be involved for the activation under strictly anoxic conditions. The involvement of a carbonylation reaction possibly leading to an aldehyde derivative is postulated for this bacterium.
7 1.9 Aim of the thesis
In the present work the mechanism of activation of acetone by the sulfate-reducing bacterium Desulfococcus biacutus is to be elucidated. It is desirable to find the product of the activation of acetone, and confirm the possible involvement of a carbonylation reaction.
Whether the initial reaction is ATP-dependent is also part of the objectives of this work.
To give a better understanding on the reaction mechanism, the enzymes that are involved in the strictly anaerobic acetone degradation by D. biacutus need to be identified.
Furthermore, obtaining the genome sequence and genome annotation of D. biacutus representing the first acetone-degrading sulfate-reducing bacterium is one of the main targets of the thesis.
8
CHAPTER 2
Carbonylation as a key reaction in anaerobic acetone activation by Desulfococcus biacutus
Olga B. Gutiérrez Acosta, Norman Hardt and Bernhard Schink
Published in Applied and Environmental Microbiology (2013) 79(20):6228-6235
2.1 Abstract
Acetone is activated by aerobic and nitrate-reducing bacteria via an ATP-dependent carboxylation reaction to form acetoacetate as the first reaction product. In the activation of acetone by sulfate-reducing bacteria, acetoacetate has not been found as an intermediate. Here, we present evidence of a carbonylation reaction as the initial step in the activation of acetone by the strictly anaerobic sulfate reducer Desulfococcus biacutus. In cell suspension experiments, CO was found to be a far better co-substrate for acetone activation than CO2. The hypothetical reaction product acetoacetaldehyde is extremely reactive and could not be identified as a free intermediate. However, acetoacetaldehyde dinitrophenylhydrazone was detected by mass spectrometry in cell-free extract experiments as a reaction product of acetone, CO, and dinitrophenylhydrazine. In a similar assay, 2-amino-4-methylpyrimidine was formed as product of a reaction between acetoacetaldehyde and guanidine. The reaction depended on ATP as a co-substrate.
Moreover, activity of aldehyde dehydrogenase (CoA acylating) tested with the putative physiological substrate was found at specific activity of 153 ± 36 mU mg-1 protein, and was specifically induced in cell-free extracts of acetone-grown cells. Moreover, acetoacetyl-CoA was detected (by mass spectrometry) after the carbonylation reaction as the subsequent intermediate after acetoacetaldehyde is formed. These results together
9
provide evidence that acetoacetaldehyde is an intermediate in the activation of acetone by sulfate-reducing bacteria.
2.2 Introduction
Acetone is produced by bacterial fermentations, for example by several Clostridium species (Han et al., 2011). It is also produced in chemistry as a solvent and as an intermediate in synthetic chemical industry. Aerobic degradation of methyl ketones was first observed with hydrocarbon-utilizing bacteria (Lukins and Foster, 1963). Acetone is degraded by some aerobic bacteria (Taylor et al., 1980) and mammalian liver cells via oxygenase- dependent hydroxylation to acetol (Landau and Brunengraber, 1987). Carboxylation of acetone to acetoacetate as a means of acetone activation was first proposed for a methanogenic enrichment culture (Platen and Schink, 1987). Requirement of CO2 as a co-substrate for acetone degradation was also observed with the nitrate reducer Thiosphaera pantotropha (Bonnet-Smits et al., 1988) and with Rhodobacter capsulatus and other phototrophs (Birks and Kelly, 1997). The reaction was studied with the nitrate-reducing strain Bun N under anoxic conditions, and it was concluded that acetoacetate was formed in an ATP-dependent carboxylation of acetone (Platen and Schink, 1989; Platen and Schink, 1990).
Attempts to measure an in vitro carboxylation of acetone at that time were unsuccessful.
However, exchange of radioactively labeled CO2 with the carboxyl group of acetoacetate was catalyzed by cell-free extracts of strain Bun N (Janssen and Schink, 1995a). A similar CO2- and ATP-dependent activation reaction was observed with the aerobic bacterium Xanthobacter autotrophicus strain Py2 (Sluis et al., 1996). A comparison between the acetone carboxylase of strain Py2 and the carboxylase of the phototrophic bacterium Rhodobacter capsulatus showed that they are identical in subunit composition (α2β2γ2 multimers of 85-, 78-, and 20-kDa subunits) and in kinetic properties (Sluis and Ensign, 1997; Sluis et al., 2002). A similar subunit composition was found recently with the acetone carboxylase of the nitrate reducer Aromatoleum aromaticum (Schühle and Heider, 2012), and with the
10
acetone carboxylases of Alicycliphilus denitrificans, Paracoccus denitrificans, and Paracoccus pantotrophus (Dullius et al., 2011). Thus, it appears well established that aerobic and nitrate-reducing bacteria activate acetone by an ATP-dependent carboxylation reaction.
Because the γ and β phosphodiester bonds of ATP need to be hydrolyzed during the reaction, two ATP equivalents are invested into a reaction that theoretically would require less than one ATP (acetone + CO2 acetoacetate- + H+ ;∆G0’ = +17.1 kJ mol-1). At least one further ATP is required for acetoacetate activation to acetoacetyl-CoA. This energy expenditure can be afforded by aerobic and nitrate-reducing bacteria because the subsequent oxidation of the acetyl moieties releases sufficient energy.
Acetone degradation by sulfate-reducing bacteria (SRB) is energetically more difficult.
Oxidation of the acetyl residue of acetyl-CoA through the CO dehydrogenase (“Wood- Ljungdahl”) pathway can form only about one ATP equivalent per acetyl residue. Thus, acetone degradation through the carboxylation reaction described above could not be supported through the subsequent oxidation of the acetyl residues. Therefore, a different mechanism for CO2-dependent acetone activation has to be postulated for these bacteria.
Acetone degradation was studied with the sulfate-reducing bacteria Desulfococcus biacutus and Desulfobacterium cetonicum (Janssen and Schink, 1995b; Janssen and Schnik, 1995). No acetone-carboxylating or acetoacetate-decarboxylating activity could be found in cell-free extracts of these bacteria. There was high acetoacetyl-CoA thiolase activity present in acetone-grown cells, but no activity of an acetoacetate-activating CoA transferase or CoA ligase. Moreover, these bacteria excreted acetate at a 1:1 ratio during growth on butyrate or 3-hydroxybutyrate, but did not accumulate acetate during growth on acetone. From these results we concluded that acetoacetate is not a free intermediate in acetone metabolism, and that activation of acetone may lead directly to an activated acetoacetyl residue, e.g., acetoacetyl-CoA (Janssen and Schink, 1995b).
Since both sulfate reducers oxidize acetyl residues through the Wood-Ljungdahl pathway, they have CO dehydrogenase activity. Therefore, they could convert CO2 to CO and employ this as a co-substrate in acetone activation, to form acetoacetaldehyde rather than
11
acetoacetate as a reaction product. In the present study, we elucidated this hypothesis with D. biacutus and found strong evidence for this novel type of reaction.
2.3 Experimental procedures Bacterial growth conditions
Desulfococcus biacutus strain KMRActS was grown in freshwater mineral medium as described before (Janssen and Schink, 1995b; Widdel and Pfennig, 1981). The medium was reduced with 1 mM sulfide, buffered with CO2/bicarbonate, and adjusted to a final pH of 7.2. Cells were grown in 1 l flasks with medium supplemented with 5 mM acetone or 5 mM butyrate as sole carbon source, and 10 mM sulfate as the electron acceptor.
Cultures were incubated under a strictly anoxic N2/CO2 (80/20) atmosphere at 30°C in the dark.
Cell suspension experiments
Cells were harvested in the late exponential growth phase at an optical density (OD 600) of 0.3. All experiments with cell extract and cell suspensions were done under strictly anoxic conditions inside an anoxic glove box. Cells were centrifuged at 6,000 x g at 10°C. The pellet was washed at least twice with 50 mM potassium phosphate (KP) buffer, pH 7.2, supplemented with 3 mM dithioerythritol as reducing agent. Cells were re-suspended in the same buffer with the addition of NaCl (1.0 g * l-1) plus MgCl2 * 6 H2O (0.6 g * l-1).
Cell suspensions with a final OD 600 of 12 were prepared in 5 ml flasks containing KP buffer with 5 mM acetone and 10 mM sulfate. The sulfate-reducing activity was measured at different time intervals for several hours. The gas phase was either N2/CO (90/10), N2/CO2 (80/20), or N2.
Preparation of cell-free extracts
Cells were harvested as described above, however, at 4°C. The cell pellet was re- suspended in the KP buffer described above, containing 0.5 mg DNase ml-1 and 1 mg ml-1 of complete protease inhibitor cocktail (Complete Mini, EDTA-free protease
12
inhibitor cocktail tablets, Roche Diagnostics GmbH, Mannheim, Germany). Cells were disrupted by passing them two times through a cooled French pressure cell at 100 MPa.
Cell debris and unopened cells were removed by centrifugation at 27,000 x g for 20 min at 4°C.
Carbon monoxide dehydrogenase (CODH) assay
Activity of CO dehydrogenase was measured at 30°C with a photometer 100-40 (Hitachi, Tokyo, Japan). Cell-free extracts of acetone-grown cells were used for enzyme assays.
Enzyme activity was tested in the already described KP buffer with the addition of 2 mM benzyl viologen (BV) as the electron acceptor. The activity was tested in cuvettes previously flushed with CO, or by addition of CO to the complete reaction mixture. The effect of CODH inhibition by potassium cyanide (KCN) was checked with a final concentration of 3 and 5 mM KCN. Reduction of BV was followed at 578 nm (ε 578 nm = 8.65 mM-1cm-1). One unit was defined as 1 µmol of BV reduced per min.
Aldehyde dehydrogenase (CoA acylating) assay
Activity of aldehyde dehydrogenase was measured in anoxic cuvettes in the same Hitachi photometer. Cell-free extracts of acetone-grown cells were used for enzyme assays;
control experiments with cell-free extracts prepared from butyrate-grown cells were run under the same conditions. Enzyme activity was followed in 50 mM KP buffer, pH 7.2, supplemented with 3 mM dithioerythritol as described before, with the addition of 2 mM coenzyme A (CoA) and 5 mM NAD+ as the electron acceptor. The reaction was started by addition of 2 mM acetaldehyde, or by addition of 20 µl of acetoacetaldehyde- containing solution (see below). NADH formation was followed at 340 nm (ε340 nm = 6.292 mM-1·cm-1). Control assays were run with boiled cell-free extracts. One unit was defined as 1 µmol of NAD reduced per min. Preparation of acetoacetaldehyde solution was done as follows: 20 µl (9.96 mg) of acetylacetaldehyde dimethyl acetal (4,4- dimethoxy-2-butanone, ALDRICH Chemistry, SIGMA-ALDRICH) was mixed with 40 µl of 37% HCl in 2 ml KP buffer, and stirred for 20 min. The reaction mix was diluted with four volumes of the same KP buffer (200 µl in 1 ml), and from this final mixture 20 to 25 µl was added to the cuvette for assay of acetoacetaldehyde dehydrogenase.
13 Activation of acetone in cell-free extract
Cell-free extracts of D. biacutus cells grown with acetone were used for enzyme assays;
control experiments were run with extracts of butyrate-grown cells. All assays were carried out under strictly anoxic conditions at 30°C. Activation of acetone was tested in a total volume of 4 ml with 5 mM acetone, 5 mM ATP, and CO (10% in the headspace) as co-substrate. The reaction mix was incubated under mild stirring for at least 3 h, and samples were taken at different time intervals with syringes previously flushed with N2. Increment of carbonyl groups was quantified with 2,4-dinitrophenyl hydrazine (DNPH) or by derivatization of the reaction product with guanidine hydrochloride to form 2- amino-4-metylpyrimidine (see methods section below). The acetone activation reaction was also tested in the presence of 5 mM potassium cyanide (KCN), an inhibitor of CO oxidation by CO dehydrogenase (Wang et al., 2013). In a further reaction setup, the same reaction mix received in addition 2 mM CoA and 5 mM NAD+. Samples of 250 µl were taken at different time intervals and acidified with 50 µl of 3 M HCl, followed by centrifugation at 10,000 x g for 10 min. The supernatant was mixed with acetonitrile (50:50) and used for the assay of acetoacetyl-CoA by electrospray ionization mass spectrometry (ESI-MS). Authentic acetoacetyl-CoA (Sigma) was used as reference.
Preparation of acetoacetaldehyde for derivatization with DNPH
Acetoacetaldehyde was prepared by chemical deprotection of acetylacetaldehyde dimethyl acetal. 9.96 mg of the protected compound was mixed with 100 µl of 1.25 M HCl in methanol, and 800 µl of acetonitrile. The reaction mix was stirred under N2 atmosphere at room temperature, and was monitored by thin layer chromatography (TLC).
Chemical synthesis of DNPH derivatives
Dinitrophenylhydrazine (DNPH; 10 mg) was dissolved in 10 ml acetonitrile or ethyl acetate, and stirred for approx. 30 min under N2 atmosphere until a clear red solution was obtained. The product of the acetoacetaldehyde dimethylacetal deprotection reaction was transferred immediately into the DNPH solution and kept at room temperature while stirring for 60 min. The reaction was followed by TLC. The product of the derivatization was purified by column chromatography using a mixture of 95% dichloromethane and
14
5% methanol as eluent. The product was analysed with ESI-MS and proton nuclear magnetic resonance (1H NMR) spectroscopy.
Isolation and characterization of DNPH derivatives from the enzyme reaction
The product of acetone activation was derivatized with DNPH. For that purpose, 300 µl of each sample taken from the reaction mix was introduced slowly into 300 µl of freshly prepared DNPH solution, and mixed for 1 h. DNPH derivatives were extracted by mixing a defined volume of the derivatization reaction mix with ethyl acetate. Derivatives were detected by high pressure liquid chromatography (HPLC), UV-spectrophotometry, and TLC. The main spot observed was scraped from the TLC plate and dissolved in dichloromethane. Further analysis using ESI-MS and 1H NMR spectroscopy was performed to characterize the derivatization product.
Derivatization with guanidine
The product of acetone activation was also derivatized with guanidine hydrochloride. The reaction conditions were set according to a procedure proposed before (Haley and Maitland, 1951). 300 µl of each sample taken from the reaction mix was introduced slowly into 500 µl of an aqueous 0.5 M guanidine hydrochloride solution (pH 9.0). The reaction was stirred for at least 24 h at 30°C. The reaction product was analysed with RP- HPLC and compared with a 2-amino-4-methylpyrimidine reference compound (ALDRICH Chemistry, SIGMA-ALDRICH).
Analytical methods
DNPH solution for quantification of carbonyl groups was prepared as follows: 0.1 g of DNPH (60% w/w) was slowly introduced into a solution of 2 M HCl. The solution was stirred for 2 h at room temperature and passed through a cellulose acetate membrane filter (Whatman ® OE 66, 0.2 µm). This solution was prepared freshly every time that it was required, as well as the standards for calibration curves. For determination of carbonyl compounds, samples from the acetone activation assay mix were slowly introduced into 100 µl of DNPH solution with further addition of 500 µl water and 100 µl 10 M NaOH. Samples were mixed for approx. 1 h, and subsequently the absorbance
15
of the hydrazone derivative was measured spectrophotometrically at 540 nm and 360 nm.
Standards of acetone dinitrophenylhydrazone were prepared the same way, and used for quantification of carbonyl groups. Analysis of 2-amino-4-methyl pyrimidine was performed by reversed-phase HPLC. A Shimadzu HPLC system equipped with UV/Vis diode array detector was used. For analysis, 50 µl samples were injected on a C-18 reverse-phase column (Grom-Sil 120 ODS, 5 µm, 150 x 4.6 mm; Grom). Eluents contained 10 mM Na2HPO4/KH2PO4 buffer, pH 7.0 (A) and acetonitrile (B), at a flow rate of 0.8 ml min−1. The elution cycle proceeded as follows: 20% B for 2 min, then a linear increase to 90% B within 9 min, and return to 20% B within 1 min, followed by an equilibration step at 20% B for 6 min. The DNPH derivative was detected with the same HPLC system using the following elution cycle: 10% B in the first minute, then a linear increase to 90 % B within 55 min, and a final 5 min equilibration at 10% B. Sulfide formed in the cell suspension experiments was quantified with the methylene blue method (Cline, 1969). Protein content of cell-free extracts was determined with the bicinchoninic acid assay (BCA protein assay kit, Thermo, Scientific). TLC was done on silica plates (silica gel 60, Merck). After drying the samples under air for 2 min, the run started with a mobile phase of 95% dichloromethane plus 5% methanol. Spots were visualized by UV light and I2 vapour. Mass spectrometric analysis was performed with an ESI source (ESI-IT: Bruker Esquire 3000 plus) in the positive and negative ion mode under the following fixed instrument settings: spray ion voltage, 1000 V; nebulizer, 13 psi; gas flow, 7 l min-1; capillary temperature, 300°C. For NMR analysis a Bruker Avance III 400 MHz spectrometer and Bruker AVIII 600 MHz spectrometer were used. 1H chemical shifts are reported relative to the residual solvent peak and are given in ppm (δ).
Spectra were measured at approx. 17°C and processed using MestReNOVA (v5.3.1).
Chemicals
Most chemicals were of analytical grade and purchased from Acros, Fluka, Sigma, Merck or Aldrich, and were used without any further purification. Dry solvents were purchased from Fluka; solvents for column chromatography were either distilled from technical grade (dichloromethane) or purchased as for chromatography grade (ethyl acetate, and methanol).
16 2.4 Results
Acetone degradation in cell suspensions
As a first approach to examine the hypothesis of possible acetone carbonylation, we checked for acetone-dependent sulfate reduction in suspensions of intact cells of D.
biacutus with CO, CO2 or N2 in the gas phase (Fig. 1A). Sulfide formation was measured as an indicator of acetone degradation. Fig. 1A shows that the highest activity and the highest extent of sulfide formation were detected with CO in the gas phase. With CO as co-substrate, sulfide was formed to a concentration of 8.5 mM after 3 h of reaction. With CO2 only about one fourth of this activity was observed, and nearly no sulfide was produced in the absence of either CO, CO2, or acetone (Fig. 1A).
Table 1. Activity of CO dehydrogenase measured in acetone-grown cell extracts of Desulfococcus biacutus
Growth condition Sp act
(mU/mg protein)
Approx.
% activity
Without KCN 882 ± 191 100
With KCN (3 mM) 121 ± 16 14
With KCN (5 mM), 10-min preincubation 24 ± 2 3 With KCN (5 mM), 20-min preincubation 13 ± 2 1
Acetone activation in cell-free extracts
Activation of acetone with CO was tested in cell-free extracts. Since we expected formation of acetoacetaldehyde as reaction product the activity was measured by quantifying keto and aldehyde groups with DNPH. In the presence of acetone, CO and ATP, cell-free extracts of D. biacutus catalyzed the formation of ketone equivalents as it is shown in Fig. 1B. Inhibition of CODH with KCN was checked before testing the
17
carbonylation reaction. According to the results in Table 1, CODH is strongly inhibited after 20 min of incubation with 5 mM KCN. Therefore, to prevent a possible oxidation of CO by CODH, KCN was added to the acetone activation reaction mix to a final concentration of 5 mM and this mix was pre-incubated for 20 min before addition of acetone. Figure 1B shows that the presence of KCN did not affect the acetone activation reaction that was measured with quantification of carbonyl groups. The reaction was stimulated by the presence of NH4+ ions or, less efficiently, by K+ ions.
Figure 1. Acetone degradation in cell suspensions and in cell-free extracts of Desulfococcus biacutus.
(A) Sulfide production in cell suspension experiments. Acetone and sulfate were added at concentrations of 5 and 10 mM, respectively, and CO and CO2 were present at initial concentration of 10 and 20% (v/v), respectively. (B) Formation of ketone equivalents measured with DNPH during acetone activation in cell-free extracts. Acetone and ATP were added at initial concentration of 5 mM each one, and CO was present at an initial concentration of 10% in the headspace. Inhibition of CO dehydrogenase was performed using 5 mM KCN. Before the addition of acetone, both samples were pre-incubated for 20 min with or without KCN.
18
Carbonylation of acetone was also tested in the same reaction system containing acetone, CO, and ATP with the addition of CoA, and NAD+. The formation of acetoacetyl-CoA was analysed by ESI-MS. In the mass spectrum shown in Fig. 2, a specific peak signal at m/z 343.6 (I) was assigned to a fraction of acetoacetyl-CoA, after the loss of 507.0 Da (II). Compared to an acetoacetyl-CoA standard, the same loss of the 507.0 Da (II) fraction was observed. The minor deviation of the mass analysis (0.5 mass units) is due to a calibration error of the ESI-MS system. The cleavage at one of the phosphorus-oxygen bonds produces the lost fraction which corresponds to 3’-Phospho-ADP. This loss has been observed to be a common phenomenon of acyl-CoA compounds (Haynes et al., 2008; Magnes et al., 2005; Norwood et al., 1990).
Figure 2. ESI-MS of acetoacetyl-CoA that was formed after the acetone activation reaction.
Activation of acetone was tested in a total volume of 4 ml with 5 mM acetone, 5 mM ATP, and CO (10% in the headspace) as co-substrate, supplemented with 2 mM CoA, and 5 mM NAD+. The CoA derivative was detected in the negative mode. The signal at m/z 343.6 (indicated as I) was assigned to a fraction of acetoacetyl-CoA, after the loss of a fractal of 507.0 Da, corresponding to 3’-Phospho-ADP (indicated as II). The signal at m/z 765.6 belongs to CoA (indicated as III).
19
Identification of products from deprotection of acetoacetaldehyde dimethylacetal
The hypothetical reaction intermediate of acetone carbonylation, acetoacetaldehyde, is known to be highly reactive (Dolgikh et al., 1964; Price, 1992) and is not commercially available. The commercially available acetoacetaldehyde dimethylacetal could be easily deprotected in acidic solution and converted to acetoacetaldehyde. While TLC analysis indicated that only one compound was formed, in the ESI mass spectrum of this deprotection reaction (Fig. 3A), a specific peak at m/z 205.2 (IV) was observed, and was attributed to the trimer of acetoacetaldehyde; a second peak at m/z 273.2 (V) was attributed to the tetramer of acetoacetaldehyde, and a third one at m/z 291.3 (VI) belonging to the tetramer of acetoacetaldehyde plus water was also observed. With 1H NMR analysis we confirmed that one of the produced compounds was triacetylbenzene (see 1H NMR analysis data). Trimerization of acetoacetaldehyde to triacetylbenzene has been observed before (Dolgikh et al., 1964; Price, 1992). Therefore, we conclude that acetoacetaldehyde was produced during the deprotection reaction and reacted with itself to form triacetylbenzene and also the tetrameric derivative. Attempts to derivatize acetoacetaldehyde with DNPH in a two-step process including deprotection and derivatization or, alternatively, in a continuous reaction, led to compounds with the specific signals of the trimer and tetramer of acetoacetaldehyde, and presented a mass spectrum pattern that was highly similar to that produced without addition of DNPH (Fig. 3B), indicating that the DNPH adduct was not formed under these conditions.
20
Figure 3. Identification of products of acetoacetaldehyde dimethyl acetal deprotection reaction.
(A) ESI-MS spectrum of the chemically deprotected acetoacetaldehyde dimethyl acetal. (B) ESI- MS spectrum after derivatization of the reaction products with DNPH. Products of acetoacetaldehyde cyclization were identified as follows: compound IV correspond to triacetylbenzene (trimmer of acetoacetaldehyde), compound V to the tetramer of acetoacetaldehyde, and compound VI represents compound V plus water.
Identification of the DNPH derivative formed during the acetone activation reaction
In order to identify the hypothetical intermediate acetoacetaldehyde, the product of acetone metabolism in cell-free extracts was derivatized with DNPH as described above.
Samples taken from the reaction mix for acetone activation in cell-free extracts were reacted with DNPH. In the ESI-MS analysis the acetoacetaldehyde-DNPH derivative ion [M+Na]+ could be observed at m/z 288.9 (VII) after 180 min of enzyme reaction (Fig.
4A). All attempts to purify the acetoacetaldehyde DNPH derivative failed because the compound proved to be unstable during the isolation process. Interestingly, after column
21
chromatography (silica column; see materials and methods) we observed in the mass spectrum the presence of the tetramer of acetoacetaldehyde peak signal at m/z 273.2 and the remains of acetoacetaldehyde-DNPH at m/z 289.2 (Fig. 4B). Unfortunately, the acetoacetaldehyde DNPH derivative that was formed after the acetone activation reaction could not be isolated for analysis by 1H NMR spectroscopy.
Figure 4. Identification of reaction products of acetone activation by cell-free extracts. (A) ESI- MS spectrum of DNPH derivative from enzyme reaction product. Compound VII was attributed to the acetoacetaldehyde DNPH. (B) ESI-MS spectrum of DNPH derivative after column chromatography. Compound V represents the tetramer of acetoacetaldehyde, and compound VII was assigned to the acetoacetaldehyde DNPH. Spectra were measured in the positive mode.
NMR analysis
The products formed during deprotection of acetoacetaldehyde dimethylacetal were analysed by 1H NMR, and resulted in the following chemical shifts: 1H NMR (400 MHz, DMSO) δ 8.62 (s, 3H), 2.72 (s, 9H). This analysis showed the presence of
22
triacetylbenzene. The spectrum of commercial triacetylbenzene (TCI Europe) was checked for comparison, and resulted in the following chemical shifts: 1H NMR (400 MHz, DMSO) δ 8.63 (s, 3H), 2.72 (s, 9H).
Derivatization with guanidine
Another strategy to trap the hypothetical acetoacetaldehyde was an N-C-N condensation reaction to form 2-amino-4-methylpyrimidine (Haley and Maitland, 1951). We employed this type of condensation reaction as well to derivatize the acetoacetaldehyde hypothetically formed in the enzymatic acetone activation reaction. Samples taken from the reaction mix were reacted with guanidine hydrochloride, and the products were analysed by RP-HPLC. The analysis showed that this product appeared exactly at the same retention time (3.65 min) as a commercial reference of 2-amino-4- methylpyrimidine, and that it presented also the same UV-absorption spectra, thus indicating that the expected reaction between acetoacetaldehyde and guanidine had occurred (Fig. 5; A and B). The formed pyrimidine could be detected only if guanidine was present at high excess. Due to the high background of excess guanidine, it was not possible to follow this reaction by ESI-MS and NMR spectroscopy. However, since we detected 2-amino-4-methylpyrimidine, it is highly probable that acetoacetaldehyde was produced during the reaction of acetone with CO, and this conclusion is also supported by the detection of the acetoacetaldehyde DNPH derivative by mass spectrometry and by the detection of acetoacetyl-CoA after the carbonylation reaction. Formation of 2-amino- 4-methylpyrimidine in the assay system described above required ATP as a co-substrate.
In the absence of ATP, no such product was formed (Fig. 6). Dependence on ATP was also confirmed in the test system using DNPH as trapping agent. The described reactions were observed only with cell-free extracts of acetone-grown cells. Control experiments with extracts of butyrate-grown cells did not produce DNPH or guanidine-reactive products.
23
Figure 5. Identification of the reaction product of acetone activation after derivatization with guanidine. (A) HPLC analysis of the 2-amino-4-methylpyrimidine reference compound and of the product formed during the reaction between the intermediate (acetoacetaldehyde) and guanidine. (B) Absorption spectra of the commercial 2-amino-4-methylpyrimidine and the product formed after the reaction between acetoacetaldehyde and guanidine.
Figure 6. Dependence of acetoacetaldehyde formation from acetone and CO on the presence of ATP as a co-substrate. The formation of 2-amino-4-methylpyrimidine was quantified as the product of the reaction between acetoacetaldehyde and guanidine. Acetone and ATP were added at initial concentrations of 5 mM. CO was present at an initial concentration of 10% in the headspace. Quantification of 2-amino-4-methylpyrimidine was done by measuring the absorption at 290nm.
24
Aldehyde dehydrogenase (CoA acylating) activity
Activity of aldehyde dehydrogenase was measured in cell-free extract of D. biacutus with acetaldehyde, and with acetoacetaldehyde that was prepared by deprotection of acetoacetaldehyde dimethylacetal. Previous experiments showed the instability of acetoacetaldehyde, nevertheless, this compound must be an intermediate before the trimer and tetramer are formed. Therefore this acidic solution was added also as a substrate for aldehyde dehydrogenase with the addition of CoA, and NAD+ as the electron acceptor. The activity was detected in cell-free extract of acetone-grown cells at 18 ± 3 mU mg-1 protein with acetaldehyde, and 153 ± 36 mU mg-1 protein with the acetoacetaldehyde preparation (Table 2).Addition of CoA caused an increase of the activity from 30 to 100%. A control assay with extract of butyrate-grown cells indicated that aldehyde dehydrogenase is specifically induced during the metabolism of acetone.
The activity with acetaldehyde increased 5 fold after addition of 20 mM NH4+ to the reaction mix.
Table 2. Aldehyde dehydrogenase (CoA acylating) activity measured in cell-free extracts of Desulfococcus biacutus.
Specific activity (mU per mg protein)
With acetaldehyde ªWith acetoacetaldehyde (-) NH4+ (+) NH4+
Acetone-grown cell-free extract 5 ± 0.5 18 ± 3 153 ± 36 Butyrate-grown cell-free extract *ND 1± 0.2 20 ± 7
* ND, Not detected
ª Ammonium addition did not stimulate the reaction with acetoacetaldehyde.
25 2.5 Discussion
In the present study, degradation of acetone under strictly anoxic conditions was investigated with the sulfate-reducing bacterium Desulfococcus biacutus. Based on our experimental results, we propose that acetone is activated by carbonylation with CO to form acetoacetaldehyde, rather than by carboxylation to acetoacetate as described for aerobic or nitrate-reducing bacteria. CO proved to be a far better co-substrate for acetone degradation than CO2 in cell suspension experiments. Since D. biacutus can reduce CO2 to CO by its carbon monoxide dehydrogenase enzyme (Janssen and Schink, 1995b) CO is available as a co-substrate for this activation reaction.
Derivatization of carbonyl compounds with DNPH has been used in the quantification of aldehydes and ketones (Cirera-Domenech et al., 2013; Huang et al., 2007; Uchiyama et al., 2011; Van den Bergh et al., 2004). In our study, the increase of ketone equivalents measured with DNPH in cell-free extracts indicated that CO and acetone were condensed to an aldehyde molecule.
A simultaneous experiment in which CO dehydrogenase was inhibited by KCN gave a similar increase of the ketone equivalents, suggesting that CO is the real co-substrate for acetone activating reaction, rather than being oxidised to CO2 by CO dehydrogenase.
This result is supported by cell suspension experiments. In control assays with chemically prepared acetoacetaldehyde, no reaction with DNPH was observed. Obviously, the formed acetoacetaldehyde had undergone a cyclization reaction to form a trimer as observed before (Dolgikh et al., 1964; Price, 1992). The formed 1,3,5-triacetylbenzene which was identified by ESI-MS and 1H NMR spectroscopy, further reacted to form the tetrameric compound. Interestingly, after the enzymatic acetone activation reaction the DNPH-acetoacetaldehyde derivative was detected by mass spectrometry with a peak at m/z 288.9, which strongly suggests that acetoacetaldehyde was indeed produced in the enzymatic activating reaction. While trying to purify this derivative we could detect the tetramer of acetoacetaldehyde (m/z 273.2) and minor amounts of the DNPH- acetoacetaldehyde derivative (m/z 289.2), indicating that the derivative might have
26
disintegrated, perhaps due to the acidic conditions that were used during the chromatographic separation process.
Formation of acetoacetaldehyde from acetone and CO is also supported by the formation of 2-amino-4-methylpyrimidine with guanidine as co-substrate, a reaction which is very specific for the detection of 1,3-dioxo aliphatics.
A further strong indication of the formation of an aldehyde as a first reaction product in acetone activation is the presence of aldehyde dehydrogenase activity. This activity was found only in extracts of acetone-grown cells, and was substantially higher when the putative physiological substrate was added, thus indicating that an aldehyde is formed specifically during degradation of acetone, and it is highly probable that this aldehyde is our hypothetical acetoacetaldehyde. Moreover, the detection of acetoacetyl-CoA after activation of acetone in the presence of CO, ATP, CoA and NAD+ supports again the formation of acetoacetaldehyde as intermediate.
The acetone-carbonylating activity was stimulated by monovalent cations such as NH4+
or K+. The activating enzyme differs from the ketone carboxylases employed by aerobic and nitrate-reducing bacteria, which depend on the presence of divalent cations such as Mg2+ and Mn2+ (Boyd et al., 2004; Jobst et al., 2010). However, the acetone-carbonylating enzyme activity of D. biacutus was stimulated by NH4+ ions, similar to the acetone carboxylases of Cupriavidus metallidurans strain CH34 and Xanthobacter autotrophicus strain Py2 (Rosier et al., 2012; Sluis et al., 2002). Neither genomic nor proteomic analysis of acetone-grown cells of D. biacutus provided any indication of acetone carboxylases similar to those described for aerobic or nitrate-reducing acetone oxidizers (unpublished results from our lab).
The observed conversion of acetone with CO to acetoacetaldehyde required ATP as a co-substrate. Perhaps this ATP is needed to stabilize the enol tautomer of acetone in the form of acetone enolphosphate. This compound is the real substrate of carboxylation by the acetone carboxylases described in the past (Boyd and Ensign, 2005; Schühle and Heider, 2012) and may as well be the real acceptor of CO in the carbonylation reaction proposed here. Since the reaction product acetoacetaldehyde is extremely reactive it
27
appears plausible that it is not released free into the cytoplasm but is immediately oxidized further to acetoacetyl-CoA, perhaps in a multienzyme complex. After all, acetone activation and conversion to acetoacetyl-CoA through this new carbonylation pathway (Fig. 7) would require a minimum of only one ATP equivalent rather than three as in the well-described carboxylation pathway, and would therefore be much better suited for bacteria operating at a small energy budget, such as sulfate-reducing bacteria.
Thus, acetone activation is another example to demonstrate that strict anaerobes such as sulfate reducers use strategies in the degradation of comparably stable compounds that are basically different from those employed by nitrate reducers, as studies with various aromatic compounds have shown in the past (Philipp and Schink, 2012). The biochemistry of the novel acetone carbonylation reaction will be subject to further studies in our lab.
Figure 7. Acetone activation mechanism by aerobic and nitrate-reducing bacteria and the proposed novel activation by carbonylation in sulfate-reducing bacteria.
28 Acknowledgments
We thank Prof. B. T. Golding (School of Chemistry, Newcastle University), Prof. Peter Kroneck (Konstanz University), and Tobias Strittmatter (Konstanz University) for valuable discussions. We thank Ines Joachim and Martin Ehrle for practical support, Antje Wiese for media preparation, and the Konstanz Research School Chemical Biology (KoRS-CB) for the fellowship granted to Olga Brígida Gutiérrez Acosta. This work was supported by the Deutsche Forschungsgemeinschaft (DFG) through the SPP 1319 priority program.
29
CHAPTER 3
ATP and thiamine pyrophosphate dependence of acetone degradation by the sulfate-reducing bacterium Desulfococcus biacutus monitored by a fluorogenic ATP analogue
Olga B. Gutiérrez Acosta, Norman Hardt, Stephan M. Hacker, Tobias Strittmatter, Bernhard Schink, and Andreas Marx.
Submitted
3.1 Abstract
Acetone can be degraded by aerobic and anaerobic microorganisms. Studies with the strictly anaerobic sulfate-reducing bacterium Desulfococcus biacutus indicate that acetone degradation by these bacteria starts with an ATP-dependent carbonylation reaction leading to acetoacetaldehyde as first reaction product. The reaction represents the second example of a carbonylation in the biochemistry of strictly anaerobic bacteria, but the exact mechanism and dependence on cofactors is still unclear. Here we present the development of a novel Förster-resonance energy transfer-based ATP probe 1 which allows to follow the consumption of ATP in the carbonylation reaction in cell-free extracts of D. biacutus in real time. The fluorogenic ATP analogue acts as activity probe to follow nucleotide-dependent enzymatic reactions. Upon processing of the probe by an ATP-hydrolysing enzyme a specific fluorescence emission is initiated. Using this analytical approach, we found that thiamine pyrophosphate acts as a cofactor to enhance the enzymatic activity in the novel acetone carbonylation reaction.
30 3.2 Introduction
Acetone is an important solvent used in chemical industry (Sifniades et al., 2011). Its microbial degradation has been studied with aerobic and anaerobic bacteria. Both nitrate- reducing and aerobic bacteria activate acetone in an ATP-dependent carboxylation reaction which consumes two ATP equivalents and forms acetoacetate as a first reaction product (Dullius et al., 2011; Schühle and Heider, 2012; Sluis et al., 2002; Sluis et al., 1996). Desulfococcus biacutus was described as a strictly anaerobic chemoorganoheterotrophic bacterium. It can grow with various substrates as carbon source, for example acetone, and uses sulfate as electron acceptor (Platen et al., 1990).
Unlike aerobic and nitrate-reducing bacteria, D. biacutus activates acetone not by carboxylation, but uses CO in the activation reaction (Figure 1). The product of this carbonylation reaction is acetoacetaldehyde, as shown recently (Gutierrez Acosta et al., 2013b). The exact mechanism of this novel reaction is still under investigation. Due to the instability of the product acetoacetaldehyde, the activating reaction is difficult to characterize, and therefore the dependence on cofactors could not be assayed so far.
Proteomic analysis of acetone-grown cells of D. biacutus indicated that a thiamine pyrophosphate (TPP)-requiring enzyme may be involved in the activating reaction (data not published). TPP is an essential cofactor in several enzymatic reactions, including pyruvate decarboxylase (EC 4.1.1.1) (Kluger, 1987), transketolase (EC 2.2.1.1) (Lindqvist et al., 1992) and pyruvate dehydrogenase (EC 1.2.4.1) (Reed, 1974). The role of TPP in these reactions is to help in the catalysis of carbon-carbon bond-forming and bond- breaking reactions that occur adjacent to a carbonyl group (Agyei-Owusu and Leeper, 2009). Consequently, our aim was to evaluate and understand a possible involvement of TPP in the activation of acetone in cell-free extracts of D. biacutus.
In order to further investigate the novel acetone activation reaction we searched for new techniques. Recently, we introduced a conceptually novel Förster-resonance energy transfer (FRET)-based assay that monitors ATP consumption in real time (Hacker et al., 2013b). In this concept, an ATP analogue is intramolecularly equipped with a fluorescent donor at the γ- or δ-phosphate, respectively, and an acceptor attached to nucleoside part of ATP. In an intact state, the excited donor fluorophore can transfer its energy to the
31
acceptor via FRET. Upon cleavage of the probe by an enzyme, the acceptor dye is spatially separated from the donor dye and the specific fluorescence emission of the donor is restored. By this concept, the activity of enzymes can be detected at high spatiotemporal resolution. In the present study, we describe the rational design and the total synthesis of a novel fluorogenic ATP analogue 1 that was tailored for the analysis of enzymatic reactions of D. biacutus. In order to use this analogue as activity probe towards ATP-dependent enzymes, it is required to ensure its stability in cell extract. Mono γ- modified nucleotides are used usually to be converted by γ-phosphate transferases and to label specific substrates (Bagshaw, 2001; Cole and Yount, 1990; Cremo et al., 1990; Lee et al., 2009; Oiwa et al., 2000; Oiwa et al., 2003; Oiwa et al., 1998; Song et al., 2008; Wang et al., 1999). Most modifications are phosphonyl- (Arzumanov et al., 1996; Green and Pflum, 2007),phosphorester-(Bettendorff et al., 2007; Freeman et al., 2010; Knorre et al., 1976; Korlach et al., 2008; Kruse et al., 1988; Lee et al., 2009; Niyomrattanakit et al., 2011; Sood et al., 2005) or phosphoramide (Green and Pflum, 2007; Green and Pflum, 2009; Knorre et al., 1976; Kruse et al., 1988; Kumar et al., 2005; Lee et al., 2009; Pollack and Auld, 1982; Song et al., 2008; Yarbrough et al., 1979; Zinellu et al., 2010) bonds, respectively. However, Marx et. al.(Hacker et al., 2012) showed recently that phosphorester modifications are stable in a wide pH range and are therefore suited to investigate enzyme reactions. Moreover, we demonstrated that modifications at the C2-position of ATP are tolerated by a broad variety of different ATP consuming enzymes (Hacker et al., 2013a). The synthesis route was developed to be as flexible as possible in order to vary the dyes of the FRET cassettes, as shown previously (Hacker et al., 2013b). However, not all previously synthesised ATP analogues are suitable for studies performed in cell extract due to limited stability of the dyes. According to our investigations sulfoCy3 and eclipse™-quencher turned out to be a FRET cassette that is stable under such conditions. Therefore, we designed and synthesized the novel γ-sulfo-Cy3, C2-eclipse- ATP analogue 1 which opens up the use of ATP activity probes in cell extracts for the first time. The novel probe carries a sulfo-Cy3 dye at the γ-moiety of the triphosphate and an eclipse™-quencher connected to the C2-position of the nucleobase. With this probe 1 in hand we provide a tailor-made nucleotide activity probe to further investigate ATP-dependent reactions, e. g., in the metabolism of D. biacutus. Using this analytical
32
approach, we focused our research on the acetone-activating reaction by D. biacutus and checked for thiamine pyrophosphate (TPP) as a possible cofactor in this unusual novel reaction.
Figure 1. Concept of the quenched ATP probe and biological pathway of interest. (A) Modified ATP probe bearing a fluorescent donor (D) attached to the γ-phosphate group and a quencher (Q) attached to the C2-position of the nucleobase. Due to FRET no fluorescent emission of D can be detected. After cleavage of the ATP probe FRET is no longer possible and direct emission of D can be observed. (B) Reaction scheme of ATP- and CO- dependent acetone activation by Desulfococcus biacutus.
3.3 Materials and Methods Synthesis of ATP analogue
The description of the organic synthesis of the ATP analogue that was used in this work can be found in Norman Hardt doctoral dissertation, or in the supporting information of the published version of this manuscript.
Cell cultures
The sulfate-reducing bacterium Desulfococcus biacutus strain KMRActS was grown in freshwater mineral medium as described before (Janssen and Schnik, 1995). The medium was reduced with 1 mM sulfide, buffered with CO2/bicarbonate, and adjusted to a final pH of 7.2. Cells were grown in 1 L flasks with medium supplemented with 5 mM acetone or 5 mM butyrate as sole carbon source, and 10 mM sulfate as the electron acceptor.
